The lymphatic system plays a dual role in fluid transport and immune surveillance. In fluid transport, extravasated fluid and macromolecules pass into lymphatic vessels that are lined by a single layer of lymphatic endothelial cells (LECs) surrounded by an incomplete basement membrane. Fluid is transferred from these vessels into larger vessels, many of which are lined by lymphatic smooth muscle that exhibit spontaneous beating, which, in concert with the action of adjacent skeletal muscle, pumps the lymph fluid back to the venous system through the thoracic duct. A healthy adult will drain one to two liters of lymph every 24 hours.
Despite apparent similarities, lymphatic vessels are very different from blood vessels, e.g., arteries, veins, and capillaries. In order to facilitate fluid uptake, initial lymph vessels are extremely permeable, are largely devoid of a basement membrane, and lack supporting pericytes and smooth muscle cells. Further, recent studies have demonstrated that, in the post-natal setting, certain molecules are expressed by LECs but not by blood endothelial cells (BECs). These molecules include, e.g., FLT-4 (also referred to as VEGF receptor-3, or VEGFR-3), D2-40, the homeobox-containing gene Prox-1, podoplanin, and the CD44 homolog LYVE-1 (Karkkainen, et al., 2002 “Lymphatic endothelial regulation, lymphoedema, and lymph node metastasis,” Semin Cell Dev Biol 13(1): 9-18). FLT-4, the protein product of the fms-like tyrosine kinase-4 gene, specifically recognizes and is activated by VEGF-C and other ligands.
Other differences between LECs and BECs have been described (Kriehuber, et al., 2001, “Isolation and characterization of dermal lymphatic and blood endothelial cells reveal stable and functionally specialized cell lineages,” J Exp Med 194(6): 797-808, incorporated herein by reference in its entirety). For example, LECs and BECs have been reported not to be functionally interchangeable. According to Kriehuber, et al., 2001, co-cultured isolated LECs and BECs form only homotypic structures. The vessel-like structures formed in these cultures segregate into structures formed entirely of LECs and structures formed entirely of BECs. Thus, BECs and cells capable of differentiating into BECs would not necessarily be expected to function in lymphatic repair and LECs and cells capable of differentiating into LECs would not necessarily be expected to function in microvascular repair.
Furthermore, it has been reported that in the vertebrate heart coronary blood and lymph vessels are derived from different sources. (See Wilting, et al., 2007, “The Proepicardium Delivers Hemangioblasts but not Lymphangioblasts to the Developing Heart,” Developmental Biology doi:101016/j.ydbio.2007.02.026).
A number of pathological conditions exist in which the ability of the lymphatic system to transport fluid is insufficient to meet demand. This leads to tissue edema that is disfiguring, disabling, and, on occasion, life-threatening. There are thus a number of settings in which modulation of expansion or repair of the lymphatic system are clinically desirable. Acute myocardial infarction leads to increased vascular permeability, thereby increasing the amount of fluid and macromolecules in the interstitial space for removal by the lymphatic system. This edema leads to tissue injury throughout the ventricle, causing histologically visible gaps between vascular endothelial cells and activation of platelets that reduce blood vessel patency. Myocardial edema is evident in many clinical states and can be caused or worsened by cardiac surgery and myocardial infarction. Myocardial edema is also implicated in rejection following heart transplant. The consequences of edema have been studied in animal models of chronic lymphatic obstruction. These studies indicate myofibrillar disruption resulting from edema-induced separation of cardiac myocytes and formation of non-elastic scar tissue which can, in turn, lead to impaired conductance and arrythmia (Kong, et al., 2005, “Effect of cardiac lymph flow obstruction on cardiac collagen synthesis and interstitial fibrosis,” Physiol Res. 55:253-258).
Severe tissue edema is frequently present following lymph node dissection, e.g., that which occurs as a routine part of surgical therapy for breast cancer. Chemotherapy and radiotherapy can also induce or worsen edema. In addition, lymphatic regeneration is an important part of wound healing. The inclusion of lymphatic vessels within tissue-engineered constructs will be an important factor in the success of these constructs and related products (Duxbury, et al., 2004, “Lymphangiogenesis in tissue-engineered small intestine,” Transplantation 77(8): 1162-6).
Effecting lymphatic expansion by administering a ligand specific for FLT-4 has been proposed (e.g., U.S. Pat. No. 6,730,658, incorporated herein by reference in its entirety). However, cellular over-expression of the FLT-4 ligand, VEGF-C, within a repairing wound reportedly induced only transient lymphatic proliferation (Goldman, et al., 2005, “Overexpression of VEGF-C causes transient lymphatic hyperplasia but not increased lymphangiogenesis in regenerating skin,” Circ. Res. 96(11): 1193-9). Further, these studies showed that VEGF-C had no ability to induce LEC migration into the area of injury. It appears that lymph vessel generation and stabilization is a multi-factorial process for which VEGF-C (or another FLT-4 ligand) is not alone sufficient.
Another important aspect of lymphatic system biology is that the lymphatics are involved in tumor metastasis. Invasion of the lymphatic system by malignant cells is well known as a means of staging tumors. In one recent study it was shown that tumor lymph vessel density was a strong predictor of positivity at adjacent sentinel lymph nodes (Massi, et al., 2006, “Tumour lymphangiogenesis is a possible predictor of sentinel lymph node status in cutaneous melanoma: a case-control study,” J Clin Pathol 59(2): 166-73). Thus, methods of altering tumor lymphangiogenesis may be applied to the management and treatment of malignancy.
LECs isolated from human palatine tonsils were reported to form tube-like structures in vitro (Garrafa, et al., 2006, “Isolation and characterization of lymphatic microvascular endothelial cells from human tonsils.” J Cell Physiol 207(1): 107-13, incorporated herein by reference in its entirety). However, harvesting of skin and tonsil LECs in sufficiently large quantities to allow for clinical use creates harvest site morbidity. The therapeutic use of LECs harvested from tumors is limited in that there is a risk of contamination with tumor cells. Adipose tissue is well-known as a source of BECs but has not previously been recognized as a source of LECs.
Data derived from kidney transplant patients have been interpreted to suggest that circulating lymphatic progenitor cells exist (Religa, et al., 2005, “Presence of bone marrow-derived circulating progenitor endothelial cells in the newly formed lymphatic vessels,” Blood 106(13): 4184-90; Kerjaschki, et al., 2006, “Lymphatic endothelial progenitor cells contribute to de novo lymphangiogenesis in human renal transplants,” Nature Medicine 12(2): 230-4, incorporated herein by reference). Religa, et al., 2005, reported that following gender-mismatched kidney transplants, approximately 4.5% of the recipient's lymphatic endothelial nuclei were found to be donor-derived. It has been reported that these cells, while circulating, exhibited certain markers of the mononuclear phagocyte lineage, including CD45 and CD14 (Kerjaschki, et al., 2006). Two other populations have been hypothesized as lymphatics EPC candidates: the FLT-4+/CD34+ population and the CD133+/FLT-4+ (e.g., Salven, et al., 2003, “VEGFR-3 and CD133 identify a population of CD34+ lymphatic/vascular endothelial precursor cells,” Blood 101(1): 168-72). However, the frequency of these populations in human peripheral blood appears to be extremely low. It has been reported that the frequency of CD34+ cells in the blood is approximately 0.2% (Bender, et al., 1991, “Identification and comparison of CD34-positive cells and their subpopulations from normal peripheral blood and bone marrow using multicolor flow cytometry,” Blood 77(12): 2591-6), and that only 0.2%±0.1% of CD34+ cells in the normal adult blood express FLT-4 (Salven, et al., 2003). Accordingly, the frequency of CD34+/FLT-4+ cells in normal blood would be only 0.04%.
Adipose tissue contains a population of cells with the ability to differentiate into multiple cell lineages, that is frequently referred to as Adipose-tissue-Derived Stem Cells (ADSC) (Zuk, et al., 2002, “Human adipose tissue is a source of multipotent stem cells,” Mol Biol Cell 13(12): 4279-95 incorporated herein by reference in its entirety and U.S. Pat. No. 6,777,231, incorporated herein by reference in its entirety.) Katz, et al., have reported performing a low density gene expression microarray analysis of ADSC (Katz, et al. 2005, “Cell surface and transcriptional characterization of human adipose-derived adherent stromal (hADAS) cells,” Stem Cells 23(3): 412-23 incorporated herein by reference in its entirety). VEGF-C, a ligand for FLT-4, was among the many genes that this study showed to be expressed in ADSC. The frequency of ADSC within the adipose-derived cell population as a whole, as measured by generally recognized clonogenic assays (fibroblastic colony-forming unit and alkaline phosphatase expressing colony-forming unit), has been reported to be approximately 1-8% (Fraser, et al., 1992, “Proliferation of totipotent hematopoietic stem cells in vitro with retention of long-term competitive in vivo reconstituting ability,” Proc Natl Acad Sci USA 89(5): 1968-72 and U.S. Pub. No. 2003/0161816, titled “Systems and Methods for Treating Patients with Processed Lipoaspirate Cells,” both incorporated herein by reference). The need for a rich source of LECs and pre-LECs, for tissue transplantation and the treatment of lymphatic diseases and disorders is manifest.